3D Bioprinted GelMA/PEGDA Hybrid Scaffold for Establishing an In Vitro Model of Melanoma

Due to the high incidence of malignant melanoma, the establishment of in vitro models that recapitulate the tumor microenvironment is of great biological and clinical importance for tumor treatment and drug research. In this study, 3D printing technology was used to prepare GelMA/PEGDA composite scaffolds that mimic the microenvironment of human malignant melanoma cell (A375) growth and construct in vitro melanoma micro-models. The GelMA/PEGDA hybrid scaffold was tested by the mechanical property, cell live/dead assay, cell proliferation assay, cytoskeleton staining and drug loading assay. The growth of tumor cells in two- and three-dimensional culture systems and the anti-cancer effect of luteolin were evaluated using the live/dead staining method and the Cell Counting Kit-8 (CCK-8) method. The results showed a high aggregation of tumor cells on the 3D scaffold, which was suitable for long-term culture. Cytoskeleton staining and immunofluorescent protein staining were used to evaluate the degree of differentiation of tumor cells under 2D and 3D culture systems. The results indicated that 3D bioprinted scaffolds were more suitable for tumor cell expansion and differentiation, and the tumor cells were more aggressive. In addition, luteolin was time- and dose-dependent on tumor cells, and tumor cells in the 3D culture system were more resistant to the drug.

specific structure that affords the necessary support for cell proliferation, migration, differentiation, ECM deposition, and response to stimuli similar to in vivo biological systems and may reduce some of the differences between in vitro and in vivo outcomes [16]. Currently, hydrogels are considered suitable materials for the biological field, while gelatin is composed of a matrix metalloproteinase (MMP) target pattern formed by hydrolysis of mammalian collagen [17]. Gelatin methacryloyl (GelMA) has become one of the most versatile biomaterials. It was originally proposed by Van Den Bulcke et al. in 2000 [18]. Polyethylene (glycol) diacrylate (PEGDA) is a hydrogel with good hydrophilicity, and its contact angle is 70° at a concentration of 1.25% [19,20]. Moreover, its elasticity is also very good, and its compressive modulus can reach 0.75 MPa at a concentration of 30% [21]. It can gel rapidly at room temperature in the presence of blue light and photoinitiators. The mechanical properties of PEGDA can vary over a wide range, and when it is mixed with GelMA, the hydrophilicity and mechanical strength of the hydrogel can be greatly improved [22]. GelMA/PEGDA hybrid hydrogel has the characteristics of good mechanical strength, high printing resolution, and resistance to degradation, and has bioactive sequences for cell attachment and matrix metalloproteinase (MMP)-sensitive degradation sites [22,23]. Based on these properties, the interaction between in vivo cells and in vitro matrix (ECM) can be simulated by an in vitro 3D model established by GelMA and PEGDA.
In this study, GelMA/PEGDA hydrogel scaffolds mimicking the tumor cell growth microenvironment were further prepared by 3D bioprinting with appropriate concentrations of GelMA and PEGDA as materials. Then, the behavior of A375 cells in both 3D and 2D culture systems was compared. In addition, the inhibitory effect of luteolin on A375 cells on 2D and 3D scaffolds was discussed and differences in drug effects were found.

Materials and Reagents
Dulbecco's modified Eagle medium (DMEM), fetal bovine serum (FBS), horse serum (HS), trypsin-EDTA and phosphate-buffered saline (PBS) were purchased from Gibco (Singapore). A375 cells were purchased from the Cell Bank of the Chinese Academy of Sciences. Fig. 1 shows the preparation process for the 3D GelMA/PEGDA scaffold and the process of exploring the antitumor effect of luteolin on A375 cells. First, composite scaffolds were prepared using 3D bioprinting technology and tumor cells were cultured in vitro for greater precision in the drug screening. The processes primarily included the synthesis of GelMA (Fig. 1A), selection of different concentrations of ink, adjustment of printing performance, bioprinting, cross-linking, and scaffold handling. Subsequently, the scaffolds were gently rinsed several times with ultrapure water and sterilized under UV light for future cell culture. We then constructed a 3D tumor model with 3D bioprinted composite scaffolds and tested the anticancer effect of luteolin and the growth of tumor cells in different culture systems in both 2D and 3D culture systems (Fig. 1B).

Preparation of GelMA
First, 100 ml of PBS buffer was heated to 50°C in a water bath. Then, 10 g of gelatin was weighed into the PBS at 50°C and stirred until completely dissolved. Following that, 8 ml of methacrylamide was slowly added to the above dissolved solution in 16 portions at a rate of 0.5 ml/min. The reaction took place in a water bath at 50°C for 2 h. Next, 400 ml of PBS buffer was added to dilute the solution, which was then stirred for 10 min. The diluted solution was poured into the co-dialysis membrane and tied tightly. The dialysis membrane was then submerged in a beaker filled with ultrapure water and dialyzed in a 40°C water bath for 5-7 days. The solution was changed three times a day and shaken well. At the end of dialysis, the supernatant was collected by centrifuge and freeze-dried for 48 h to obtain GelMA solid [24].

Characterization of Materials
Preparation of GelMA/PEGDA scaffold. Solutions of GelMA/PGEDA with mass fractions of 2.5/2.5%, 5/ 2.5%, 7.5/2.5%, 10/2.5%, 12.5/2.5% (w/v) were prepared by dissolving GelMA and PEGDA in PBS buffer. The mixed solutions were cured and cross-linked under blue light to form a cylinder with a radius of 8 mm and a height of 5 mm.
Young's modulus test. Young's modulus of different scaffolds was evaluated by using a uniaxial compression tester (Norwood 5556 Research Institute, USA). Prior to testing Young's modulus of the scaffolds, the dry scaffolds were soaked in PBS for 24 h. The scaffolds were then removed and excess water was gently wiped from the surface with filter paper. The initial cross-sectional area and thickness of each sample were then measured. The next step was to slowly apply a preload force to the samples. The compression rate was set to 0.1 mm/min. When the scaffold was compressed to about 50% of its original length, the modulus of compression was calculated based on the slope of the initial linear region of the stress-strain curve.
Water absorption test. Three parallel samples were provided for each of the cylindrical scaffolds of different concentrations. The samples were placed in Petri dishes of known mass and 4 ml of PBS solution was added to each and soaked at 37°C for 24 h to reach the swelling equilibrium. The PBS was aspirated, the remaining water droplets were wiped off with filter paper, and the swollen sample masses (W s ) were weighed. All samples were freeze-dried and weighed to obtain their dry weight (W d ). The swelling rate (Q s ) of the samples was calculated according to the following equation: . (1) Porosity. Porosity was measured by the drainage method. The radius (r) and height (h) of each lyophilized scaffold were measured and weighed (M 1 ). The scaffolds were then immersed in absolute ethanol and placed in a vacuum drying oven with reduced pressure to remove air bubbles. After 3 h of immersion, the surface water was filtered off with filter paper and then weighed on an electronic balance (M 1 for the weight before immersion and M 2 for the weight after immersion). The porosity (θ) is calculated by the following formula (ρ is the density of ethanol 0.789 mg/ml): . (2) Scanning electron microscopy (SEM) analysis. The lyophilized scaffolds were obtained after lyophilization, and the scaffolds were cut along the cross section with a scalpel and sprayed with gold. The microstructure of the lyophilized scaffolds was observed by SEM (JEOL, Japan) at an accelerating voltage of 5V.
Cytotoxicity test. Live/dead cell staining experiments were performed to observe the viability of A375 cells cultured in GelMA/PEGDA complex gels at different concentrations. Live and dead cells were double-stained with calcein-AM and propidium iodide (PI). Calcein-AM is a cell-staining reagent that fluorescently labels live cells. When it enters the cytoplasm, esterases hydrolyze it to calcein and leave it inside the cell, emitting green fluorescence. PI reaches the nucleus only through the disordered region of the dead cell membrane and is embedded in the cell's DNA double helix structure, producing red fluorescence. The above hydrogel solutions at different concentrations were first added to 24-well plates at 0.5 ml/well, solidified with blue light, and then inoculated with 100 μl of cell suspension (5 × 10 6 cells/ml) and incubated in a 37°C, 5% CO 2 incubator for 24 h. After 1 day of culture, calcein-AM/PI double-stained working solutions were prepared and incubated in a humidified incubator for 45 min. Then, the dye solutions were aspirated and the samples were washed three times with phosphate-buffered saline (PBS). Morphological photographs of the stained cells were taken with a Cytation 5 Imaging Reader (BioTek, USA). The number of live and dead cells was analyzed from the fluorescent images.

3D Bioprinting
GelMA and PEGDA were used as materials to prepare 3D bioprinted scaffolds to build in vitro tumor models. The 3D printing ink was prepared from GelMA and PEGDA in a 4:1 by mass ratio to obtain a 10/2.5% GelMA/ PEGDA mixture. The 3D scaffolds were designed using computer-aided design software. The length, width, and height of the 3D scaffold were 10 mm, 10 mm, and 1.2 mm, respectively. The spacing of 3D supports is 0.8mm and the number of layers is 6. The 3D scaffold was then printed by an extrusion printing technique using a 3D bioprinter (EnvisionTEC GmbH, Germany). The radius of the printing needle was 0.08 mm, the print speed was set to 12.5 mm/s, the pressure was 1.7 bar, and the temperature was set to 17°C. After printing, the scaffold was shaped by irradiating it with blue light for 1 min. Then, after the scaffolds were completely cross-linked, they were sterilized under a UV sterilization lamp for 24 h. Finally, the scaffolds were gently rinsed several times with deionized water and stored for subsequent cell experiments.

Cell Culture
A375 cells were cultured in DMEM containing 10% FBS and 1% penicillin-streptomycin in a humidified incubator at 37°C and 5% CO 2 . When the cells covered 80-90% of the bottom of the culture flask, a passage experiment was performed to ensure viability for subsequent usage of the cells.

Cell Growth Activity Assay
The activity of A375 cells in 2D and 3D scaffolds was detected by the live/dead staining method and CCK-8 method, respectively. First, 3D printed scaffolds were inoculated with 100 μl of cell suspension (2 × 10 6 cells/ml). Cell scaffold complexes were placed on 12-well plates. After 1, 3, 5, and 7 days of incubation, the cells were stained with a staining solution consisting of 2 μmol/l calcein-AM and 4 μmol/l PI and incubated at 37°C with 5% CO 2 for 45 min. The dye solution was then removed with a pipette and the samples were washed three times with phosphate-buffered saline (PBS). An enzyme marker was used to detect cell viability. In the 2D scaffold, 540 μl of mixed hydrogel solution was added to each well of a 12-well plate for UV-cured cross-linking. The same number of cells were then inoculated in the 12-well plates and cultured for 1, 3, 5, and 7 days and assayed as described above. Considering the effective range of OD values, 5,000 cells were inoculated in 2D and 3D scaffolds when performing the CCK-8 analysis. After 1, 3, 5, and 7 days of cell culture, 100 μl of fresh medium was replaced with 10 μl of CCK-8 working solution and incubated in an incubator for 40 min. Then,100 μl of the test solution was added dropwise to a 96-well plate and the absorbance was measured at 450 nm with a time of 10 s. Cell proliferation and activity were compared in 2D and 3D cultures.

Phalloidin/DAPI Staining
To assess the growth of tumor cells on 2D and 3D scaffolds, a 100 μl A375 cell suspension (2 × 10 6 cells/ml) was inoculated on the scaffolds and incubated for 12 h. The culture medium was removed and washed 3 times with sterile PBS. The medium was then incubated in an incubator for 5 days using DMEM medium containing 10% FBS. The differentiated cell scaffolds were fixed with 4% paraformaldehyde (PFA) solution and infiltrated with 0.3% Triton X-100 solution. Cells attached to the scaffold were stained with phalloidin and fluorescence images were acquired using the Cytation 5 Imaging Reader (Bio-Tek).

Immunofluorescence Staining
After 5 days of differentiation culture, the scaffolds were washed 3 times with PBS. The scaffolds were fixed with 4% PFA for 15 min, then permeabilized with 0.5% Triton X-100 (PBS preparation) for 20 min at room temperature, washed with PBS, aspirated, and closed with normal goat serum for 30 min at room temperature. Then, the closure solution was aspirated, a sufficient amount of diluted primary antibody was added dropwise, and the solution was left overnight at 4°C. The next day, the cell scaffold was reacted with a secondary antibody dilution of goat anti-rabbit IgG antibody-Alexa Fluor 546 coupling (1:200, Invitrogen) for 30 min and incubated with DAPI solution for 15 min at room temperature. Fluorescent images were obtained using the Cytation 5 reader (Bio-Tek).

Cell Activity Assay After Exposure to Luteolin
Live/dead staining and CCK-8 analysis were used to assess the impact of different concentrations of luteolin on A375 cell growth. Luteolin was dissolved in the culture medium at different concentrations and decontaminated with a filter membrane (pore size 0.22 μm). Then, 2 × 10 5 A375 cells were inoculated on 2D stents and 3D printed scaffolds and cultured for 5 days to form tumor cell clusters. They were then exposed to different concentrations of luteolin for 24 and 48 h and cell viability was assessed by CCK-8 and live/dead staining assays.

Statistical Analysis
All data are expressed as mean ± standard deviation (SD). One-way analysis of variance (ANOVA) was used for significance analysis. p < 0.05 was considered a significant difference. OriginPro2018 software (Origin Lab, USA) was used for all statistical analyses.

Characterization of GelMA/PEGDA Scaffolds
Structural support for cell growth is provided by GelMA [25]. Pure gelatin lacks mechanical strength, and the shrinkage of gelatin during the manufacturing process destroys the microstructure. On the other hand, natural gelatin comes with the risk of immunogenicity and pathogen transmission. Gelatin methacrylate (GelMA) is prepared from methacrylic anhydride (MA) and gelatin (gelatin). It is a photosensitive biohydrogel material with good biocompatibility. It can generate a solidification reaction under the excitation of blue light to form a threedimensional structure with certain strength suitable for cell growth and differentiation. The biocompatibility of GelMA is much better than that of Matrigel and Fibrin Glue, and its properties are similar to collagen. At the same time, its molding properties are much better than that of collagen, making it the best choice to replace these other materials. Polyethylene glycol (ethylene glycol) diacrylate (PEGDA) hydrogels are considered bioinert, and their mechanical properties can vary over a wide range of moduli. GelMA/PEGDA hybrid hydrogel has the characteristics of good mechanical strengths, high printing resolution, and resistance to degradation. Tumor models based on 3D printing can provide cell adhesion sites, which can change the gene expression and signal transmission of tumor cells to a certain extent. In addition, the bioactive sequences for cell attachment and matrix metalloproteinase (MMP)-sensitive degradation sites are retained in the gelatin skeleton [23], which helps to research melanoma cell invasion and aggregation [26]. Furthermore, the low molecular weight of PEGDA gives it more flexibility in the reaction system. PEGDA is a bifunctional monomer that increases the number of unsaturated double bonds in the system and improves the degree of cross-linking of the hydrogel. The pure GelMA scaffold is not suitable for long-term culture of melanoma cells with rapid proliferation in a culture environment of 37°C and 5% CO 2 . Therefore, the addition of PEGDA keeps the hydrogel stable for a long time at 37°C. Both natural materials are extensively used to construct a 3D scaffold in a cell culture system. Even more encouraging, the blend of GelMA and PEGDA has also been reported to improve the bioactivity and mechanical properties of hydrogels, and due to the hydrophilic nature of PEGDA, it also helps to improve hydrophilicity as well as oxygen and nutrient exchange [23]. Considering these factors, GelMA and PEGDA are suitable materials for the construction of 3D scaffolds.
To prepare 3D hydrogel scaffolds, the GelMA/PEGDA hydrogels were made into a cylindrical solid by a UV light-curing technique. Then they are freeze-dried to obtain lyophilized scaffolds. The cross-section of the hydrogels (Fig. 2E) exhibits a loose, porous morphology, which facilitates the uptake of oxygen and nutrients. The pore size of the hydrogel material is estimated to be 129 ± 13 μm. Pore sizes that are too narrow may limit cell proliferation, but scaffolds with pore sizes that are too wide will decrease the surface area for cell adherence. This lattice structure may enhance the mechanical properties of the material and reduce the risk of scaffold lysis. In addition, this interoperability of pores conveniently provides for better nutrient exchange and cell migration [27]. In general, 3D printing ink has excellent mechanical properties and can support tumor cells in culture for long periods. The results of mechanical performance tests showed that the stress on the scaffold of the GelMA/PEGDA hybrid hydrogel becomes higher with the increase of GelMA concentration ( Fig. 2A). Taking the hydrogel with G10/P2.5 concentration as an example, the stresses of the scaffold reached 5.0311 kPa and 52.210 kPa at strains of 0.1 and 0.5, respectively, indicating that the scaffold has good compressive properties. The Young's modulus also revealed the same tendency as the compressive stress curve (Fig. 2B). In addition, the swelling rate and porosity of the hybrid hydrogel were tested. As shown in Fig. 2C, the results indicated that the hybrid hydrogel has good hydrophilicity and the water absorption performance of the material will slightly decrease with increasing GelMA concentration. The water absorption and porosity of the G10/P2.5 scaffold were 649.54 ± 45.60% and 74.93 ± 1.87% (Fig. 2D). The performance tests of mixed hydrogels at different concentrations showed that GelMA/ PEGDA could be applied to cellular experiments.
Based on these good physical properties, we further tested the biocompatibility of the hybrid hydrogel material. According to Fig. 2F, the number of A375 cells was relatively low on the hydrogel at a concentration of G2.5/P2.5. The reason may be that the hydrogels were too soft for the cells to adhere well and were washed away when washed with PBS [28]. The amount of cells in the G5/P2.5 group was relatively high, but there was still a problem with cell attachment. With the increase of GelMA concentration, the mixed hydrogel A375 cells could adhere well at the three concentrations of G7.5/P2.5, G10/P2.5 and G12.5/P2.5. The results of live/dead staining experiments showed that a large number of cells survived, indicating that the mixed hydrogel material with this concentration has good biocompatibility. Based on this, the well-performing G10/P2.5 concentration was used as a follow-up experiment for 3D bioprinting.

2D and 3D Cell Culture
Excellent cytoactive is a key requirement for the construction of in vitro tumor models. Live/dead staining and CCK-8 assays were used to explore the liveness of A375 cells on the scaffold (Fig. 3B). As shown in Fig. 3A, most living cells were stained with calcein-AM on different days and expressed strong green fluorescence. The green fluorescence was significantly enhanced as the incubation time increased, indicating that A375 cells kept up high liveness properties on the scaffold and the amount of dead cells stained by PI showed a slight upward trend. The main causes of cell death were insufficient nutrition and oxygen supply [29]. The staining results also showed that as the number of days of cell culture increased, the cells continued to proliferate until they covered the entire 3D scaffold. After 7 days of culture, the cells on the 3D scaffold could grow in aggregates on the scaffold, showing spherical cell clusters due to the structural characteristics and good biocompatibility of the 3D bioprinted scaffold. Compared with 2D scaffolds, 3D scaffolds could better promote the aggregation and growth of tumor cells. CCK-8 assays were used to detect and compare the proliferation of 2D and 3D scaffold-cultured cells (Fig. 3D). Furthermore, the results of calcein-AM and PI staining as well obviously indicated that the cells continued to proliferate during 7 days of culture. On day 1, cell proliferation was lower in the 3D scaffold than in the 2D scaffold. The difference in cell numbers on day 3 gradually decreased as the incubation time increased. From the fifth day, cell proliferation on the 3D scaffolds exceeded that on the 2D scaffolds. With fewer culturing days, the proliferation of cells on 2D scaffolds was higher than that of cells on 3D scaffolds. The reason may be related to the adaptability of the cells. For a short period, cells are more likely to attach and proliferate on a flat surface. When the culture time is longer, as the cells keep proliferating, the 2D scaffold will show cell contact inhibition faster than the scaffolds were used to assess A375 cell activity with CCK-8 after 1, 3, 5 and 7 days of culture (n = 3). ***p < 0.001, **p < 0.01, *p < 0.05. 3D scaffold, while the 3D scaffold's three-dimensional structure can provide a larger growth area for the cells, and thus the cells can proliferate more.

Phalloidin /DAPI Staining
After 5 days of inoculation culture, the 2D-cultured and 3D-cultured A375 cells were stained with FITC-labeled phalloidin and their specific morphology was observed under a microplate reader . As shown in Fig. 4, the 2Dcultured A375 cells were laid flat on a flat surface, and the staining image on the 3D scaffold showed that A375 cells spread around the scaffold to the edge, the cells in the middle part aggregated into clusters, and the A375 cells were more elongated in shape. DAPI staining results showed that the 3D scaffold had a different height spatial distribution of cells compared to the 2D scaffold, which was closer to the human environment. In different culture systems, the growth morphology of the cells on the 2D scaffold and the 3D-printed scaffold was found to be different [30]. In the 3D culture system, the 3D-cultured A375 cells showed stronger aggregation and migration. The results suggested that A375 cells clustered more easily and dispersed more fully in the 3D culture system.

Immunofluorescence Staining
Immunofluorescence staining experiments further verified the effect of different culture systems of 2D scaffolds and 3D printed scaffolds on the differentiation ability of A375 cells. MMP-9 is a matrix metalloproteinase whose main function is to degrade and remodel the extracellular matrix [31]. In addition to the prototypical structure of MMPs, the catalytic region of MMP-9 includes three repetitive fibronectin structural domains. This structural domain has a high affinity for gelatin, which fits well with the material used in our scaffold. In addition, MMP-9 can be involved in angiogenesis through the release of vascular endothelial growth factor (VEGF) [32]. It is well known that blood vessels play a very important role in tumor formation. Green fluorescence-labeled secondary antibodies were used to detect MMP-9 protein. Moreover, DAPI was used for staining to determine the location of the cell nucleus. The stronger the green fluorescence signal, the higher the differentiation of the surface cell protein. As shown in Fig. 5A, cells on 2D and 3D scaffolds have differentiated to express MMP-9 protein, and the intensity of the green fluorescence signal was significantly different in different culture systems. In addition, semiquantitative analysis of the images using ImageJ software showed that the relative fluorescence intensity of the 3D scaffolds increased by 88.24 ± 6.61% compared to the 2D scaffolds (Fig. 5B). The fluorescence area on the 3D scaffold was larger and the MMP-9 protein expression level of A375 cells was higher compared to the 2D scaffold. The results suggest that A375 cells are more aggressive in the 3D culture system and more closely resemble tumor growth in vivo.

Effect of Luteolin on A375 Cell Apoptosis
To test the inhibitory effect of luteolin on tumor cells on 2D and 3D scaffolds, A375 cells were seeded at a high density and cultured for 5 days to form a cell mass (Figs. 6A and 6B). After 5 days of culture, luteolin at gradient concentrations of 0-250 μg/ml was added and incubated for 24 and 48 h. Live/dead stained images clearly show the growth of a cell mass on the scaffolds. After the undosed group continued to be cultured for 2 days, the cells almost filled the large pores of the scaffold and spread in all directions. However, when the concentration of luteolin increased to 100 μg/ml, most of the cells were detached from the scaffold and the number of cells decreased observably (Fig. 6A). When the concentration increased to 200 μg/ml, the number of cells on the 3D scaffold also decreased significantly (Fig. 6B). We observed that the green fluorescence of the 2D and 3D scaffolds was significantly reduced after drug treatment, and the amount of live cells was also significantly reduced. With the extension of the treatment time, the staining results of the scaffolds at the same drug concentration for 48 h showed a further decrease in the amount of live cells compared to the scaffolds at 24 h. On this basis, the results comparing the drug inhibition in 2D and 3D tumor models were shown in Fig 6. In the presence of the same concentration of luteolin, A375 cells showed greater resistance in the 3D culture model than in the 2D culture model. This is because the drug sensitivity of tumor cells in the 3D tumor model is lower than that of the 2D culture model. As shown in Fig. 6A, after adding the drug to the 2D scaffold, the cells showed a dispersed monolayer distribution, and luteolin could act directly on the cells. The cells on the 3D scaffold showed an aggregated distribution, and the drug could only act directly on a portion of the cells on the surface of the cell mass. The cells inside were exposed to lower concentrations of the drug than the peripheral cells that were directly exposed to the drug. Hypoxia or cell cycle changes in the 3D tumor model may also lead to differences in drug sensitivity [27]. Another reason for this could be the continuous accumulation of A375 cells on the 3D scaffold, forming cell clusters in the pores of the scaffold, which helps to achieve cell-cell and ECM interactions that are closer to the in vivo environment [12]. On the contrary, 2D culture could not form a tumor tissue structure similar to the in vivo environment, which limits the morphology and survival of cells.
CCK-8 assay was used to compare the survival rates of A375 cells cultured in 2D and 3D scaffolds at different drug concentrations. From the results shown in Fig. 6C, it can be noted that luteolin had a significant effect on the proliferation of A375 cells, which is consistent with the results of the live/dead staining method analysis. Figure 6A showed the absorbance of A375 cells after exposure to luteolin (50, 100, 150, 200, 250 μg/ml), measured with CCK-8. In the presence of different concentrations of luteolin, the cell survival rate decreased significantly with increasing time of drug action. The reason was that the inhibitory effect of the drug was relatively small during the first 24 h. After 48 h, the effect of the drug was further enhanced, apoptosis was intensified, and the number of cells was further reduced. A375 cells showed a dose-and time-dependent response to luteolin, and the higher the concentration and the longer the drug action time, the lower the cell survival rate [33,34]. Fig. 6C also showed that the cell survival rate on 3D-printed scaffolds was higher than 2D-cultured scaffolds at the same drug concentration, which is consistent with the junction analysis described above.
Live/dead staining and CCK-8 assays showed that the inhibitory effect of luteolin on 2D-and 3D-cultured A375 cells was time-and dose-dependent. Although there were differences in drug sensitivity between the two culture systems, it was certain that luteolin has an inhibitory effect on A375 tumor cells.
In summary, the GelMA/PEGDA scaffold produced by 3D bioprinting technology provides a more efficient means for the in vitro culture of A375 tumor cells. We used a combination of extrusion printing and UV light curing. With the addition of a photoinitiator, both GelMA and PEGDA can cure quickly at 365nm light, which is conducive to the stacking of printing layers and the rapid prototyping of 3D structures. The combination of lightcuring technology improves the precision of extrusion printing [22]. The mechanical strength, porosity, swelling rate and biocompatibility of GelMA/PEGDA scaffold material were tested to prove that its performance meets the requirements of the study. SEM results showed that the porosity and pore size of the scaffold material are suitable for cell adhesion and growth. The results of CCK-8 and live/dead staining indicated that the 3D scaffold is more suitable for the proliferation of tumor cells. In addition, cell morphology staining results showed that 3D- bioprinted hydrogel scaffolds were more suitable for tumor cell expansion and differentiation compared to 2D culture. Preliminary studies of protein expression in cells on 2D and 3D scaffolds were also performed, showing that cells cultured on 3D-bioprinted scaffolds had higher levels of MMP-9 protein secretion and that the cells exhibited greater aggressiveness. Studies on the effect of luteolin on A375 cells also revealed that the cytotoxicity of luteolin was time-and concentration-dependent. Furthermore, 22.62 ± 1.92% survival was achieved in 2D culture (50 μg/ml, 48 h) and 31.81 ± 2.61% was achieved in 3D culture (50 μg/ml, 48 h). In the 3D culture system, A375 cells were more resistant to the drug. In conclusion, 3D-bioprinted cell mass models offer a new approach for the construction of in vitro tumor models and the screening of anticancer drugs with great potential for further development. (C) Survival of A375 cells exposed to luteolin was assessed by CCK-8 assay after 24 and 48 h of culture, respectively (n = 3). ***p < 0.001, **p < 0.01, *p < 0.05.


Co-Application of Statin and Flavonoids as an Effective Strategy to Reduce the Activity of Voltage-Gated Potassium Channels Kv1.3 and Induce Apoptosis in Human Leukemic T Cell Line Jurkat

Voltage-gated potassium channels of the Kv1.3 type are considered a potential new molecular target in several pathologies, including some cancer disorders and COVID-19. Lipophilic non-toxic organic inhibitors of Kv1.3 channels, such as statins and flavonoids, may have clinical applications in supporting the therapy of some cancer diseases, such as breast, pancreas, and lung cancer; melanoma; or chronic lymphocytic leukemia. This study focuses on the influence of the co-application of statins—simvastatin (SIM) or mevastatin (MEV)—with flavonoids 8-prenylnaringenin (8-PN), 6-prenylnarigenin (6-PN), xanthohumol (XANT), acacetin (ACAC), or chrysin on the activity of Kv1.3 channels, viability, and the apoptosis of cancer cells in the human T cell line Jurkat. We showed that the inhibitory effect of co-application of the statins with flavonoids was significantly more potent than the effects exerted by each compound applied alone. Combinations of simvastatin with chrysin, as well as mevastatin with 8-prenylnaringenin, seem to be the most promising. We also found that these results correlate with an increased ability of the statin–flavonoid combination to reduce viability and induce apoptosis in cancer cells compared to single compounds. Our findings suggest that the co-application of statins and flavonoids at low concentrations may increase the effectiveness and safety of cancer therapy. Thus, the simultaneous application of statins and flavonoids may be a new and promising anticancer strategy.

Introduction
Voltage-gated potassium channels of the Kv1.3 type are widely expressed in many types of cells, both in the plasma membrane and in the inner mitochondrial membrane (mito Kv1.3 channels) [1][2][3][4]. The activity of Kv1.3 channels is required, among others, in cell proliferation and apoptosis [2][3][4][5]. The specific inhibitors of Kv1.3 channels may potentially be applied in the treatment of T-lymphocyte-mediated autoimmune diseases [4,5] and chronic respiratory diseases, including severe cases of COVID-19 disease [6,7]. Since the expression of Kv1.3 channels is up-regulated in some cancer types, such as breast and lung cancer, melanoma, pancreatic ductal adenocarcinoma, or chronic lymphocytic leukemia (B-CLL) [4,[8][9][10], inhibitors of the channels may also clinical applications in supporting the chemotherapy of these types of cancer [9][10][11]. Among inhibitors of Kv1.3 channels, the most promising candidates for potential clinical application in cancer therapy are lipophilic small-molecule organic compounds that are able to induce selective cancer cell apoptosis Figure 1B depicts relative peak currents (see Materials and Methods) calculated upon co-application of the statins and each of the statins alone. The currents were equal to 0.47 ± 0.14 (n = 28), 0.50 ± 0.09 (n = 16), and 0.41 ± 0.11 (n = 21) of the control value upon coapplication of simvastatin and mevastatin and the application of simvastatin alone and mevastatin alone, respectively. The currents recorded upon a co-application of simvastatin and mevastatin and in the presence of simvastatin or mevastatin applied alone did not significantly differ one from another (p > 0.05, one-way ANOVA). Thus, a co-application of simvastatin and mevastatin does not influence the magnitude of their inhibitory effect on Kv1.3 channels.
It is known that the inhibitory effects of both studied statins are accompanied by a significant acceleration of the inactivation rate of Kv1.3 currents by means of a significant reduction in the inactivation time constant [12]. Therefore, it was of interest to prove whether a co-application of simvastatin and mevastatin could further influence the inactivation rate of the currents. Figure 1C shows normalized whole-cell potassium currents recorded applying depolarizing voltage steps (see in Materials and Methods) under control conditions upon the co-application of simvastatin and mevastatin (both applied at a concentration of 6.0 μM) and after the wash-out of the compounds. Apparently, a co-application of simvastatin and mevastatin accelerated the inactivation of the currents. The The figure depicts raw currents containing two components: small linear and much bigger non-linear currents. The linear current was the unspecific leak current, not interesting for our study, whereas the non-linear one was the actual Kv1.3 channel current [24]. This current was significantly diminished upon co-application of the statins. The current recovered completely after the wash-out of the compounds. This indicates that the inhibitory effect of the statins on the channels is reversible. Figure 1B depicts relative peak currents (see Materials and Methods) calculated upon co-application of the statins and each of the statins alone. The currents were equal to 0.47 ± 0.14 (n = 28), 0.50 ± 0.09 (n = 16), and 0.41 ± 0.11 (n = 21) of the control value upon co-application of simvastatin and mevastatin and the application of simvastatin alone and mevastatin alone, respectively. The currents recorded upon a co-application of simvastatin and mevastatin and in the presence of simvastatin or mevastatin applied alone did not significantly differ one from another (p > 0.05, one-way ANOVA). Thus, a co-application of simvastatin and mevastatin does not influence the magnitude of their inhibitory effect on Kv1.3 channels.
It is known that the inhibitory effects of both studied statins are accompanied by a significant acceleration of the inactivation rate of Kv1.3 currents by means of a significant reduction in the inactivation time constant [12]. Therefore, it was of interest to prove whether a co-application of simvastatin and mevastatin could further influence the inactivation rate of the currents. Figure 1C shows normalized whole-cell potassium currents recorded applying depolarizing voltage steps (see in Materials and Methods) under control conditions upon the co-application of simvastatin and mevastatin (both applied at a concentration of 6.0 µM) and after the wash-out of the compounds. Apparently, a co-application of simvastatin and mevastatin accelerated the inactivation of the currents. The current did not recover completely after the wash-out of the statins. This is in agreement with our earlier results, which showed that acceleration of the inactivation by simvastatin was partially irreversible [12]. Figure 1D depicts values of inactivation time constants upon co-application of the statins and each of the statins alone at the same concentration. The values were equal to 18.19 ± 3.97 ms (n = 20), 16.95 ± 4.2 ms (n = 31), and 24.56 ± 11.57 ms (n = 11) upon the co-application of simvastatin and mevastatin and upon the application of simvastatin and mevastatin alone, respectively.The inactivation time constants were not significantly different one from another (p > 0.05, one-way ANOVA). Thus, the co-application of simvastatin and mevastatin had no influence on the inactivation rate of Kv1.3 currents. Figure 2A depicts relative peak currents upon co-application of simvastatin with 8-prenylnaringenin, simvastatin applied alone, and 8-prenylnaringenin applied alone, respectively, whereas Figure 2B shows the same data obtained upon co-application of mevastatin with 8-prenylnaringenin, mevastatin applied alone, and 8-prenylnaringenin applied alone. The concentration of 8-prenylnaringenin was chosen to be 3 µM because such a concentration was high enough to inhibit most of the Kv1.3 channels in Jurkat T cells [21]. The concentrations of the statins were equal to 6 µM. The relative peak current upon co-application of simvastatin and 8-prenylnaringenin was equal to 0.12 ± 0.07 (n = 15). This value was significantly smaller than the values obtained for simvastatin (see above) and 8-prenylnaringenin applied alone (0.39 ± 0.10, (n = 15), p < 0.05, one-way ANOVA). The inactivation time constant for simvastatin co-applied with 8-prenylnaringenin was equal to 13.89 ± 4.39 (n = 12). This value was not significantly different from the values calculated for simvastatin applied alone (see above, p > 0.05, one-way ANOVA), but it was significantly smaller than the values obtained for 8-prenylnaringenin applied alone (35.52 ± 8.99 (n = 13), p < 0.05, one-way ANOVA).
The relative peak current for mevastatin co-applied with 8-prenylnaringenin was equal to 0.09 ± 0.064 (n = 19). This value was significantly smaller than the values obtained for mevastatin and 8-prenylnaringenin applied alone (see above, p < 0.05, one-way ANOVA). The inactivation time constant upon co-application of simvastatin and mevastatin with 8-prenylnaringenin was not significantly different from the value calculated for simvastatin and mevastatin applied alone (data not shown). Figure 2C depicts relative peak currents upon co-application of simvastatin with 6-prenylnaringenin, simvastatin applied alone, and 6-prenylnaringenin applied alone, respectively, whereas Figure 2D shows the same parameter obtained upon co-application of mevastatin with 6-prenylnaringenin, mevastatin applied alone, and 6-prenylnaringenin applied alone. The concentration of 6-prenylnaringenin was equal to 6.0 µM because such a concentration was high enough to significantly inhibit Kv1.3 channels in Jurkat T cells [19]. The concentrations of the statins were previously mentioned. The relative peak current for simvastatin co-applied with 6-prenylnaringenin was equal to 0.26 ± 0.17 (n = 18). This value was significantly smaller than the values obtained for simvastatin (see above) and 6-prenylnaringenin applied alone (0.79 ± 0.09 (n = 11), p < 0.05, one-way ANOVA). The inactivation time constant for simvastatin co-applied with 6-prenylnaringenin was equal to 18.9 ± 6.31 (n = 14). This value was not significantly different from the values calculated for simvastatin applied alone (see above, p > 0.05, one-way ANOVA), but it was significantly smaller than the value obtained for 6-prenylnaringenin applied alone (68.33 ± 20.74 (n = 3), p < 0.05, one-way ANOVA).
The relative peak current for mevastatin co-applied with 6-prenylnaringenin was equal to 0.29 ± 0.069 (n = 20). This current was significantly smaller than the currents obtained for mevastatin and 6-prenylnaringenin applied alone (see above, p < 0.05, one-way ANOVA). The inactivation time constant for simvastatin and mevastatin co-applied with 6-prenylnaringenin was not significantly different from the value calculated for simvastatin and mevastatin applied alone (data not shown).  The fact that the magnitude of channel inhibition upon co-application of the statins with selected flavonoids is significantly stronger than the effect exerted of each compound applied alone may be a consequence of the additivity of their inhibitory effects. If two channel inhibitors act on the same molecular target independently of each other, their inhibitory effects may be additive. In such a case, the relative peak current upon co-application of the inhibitors is equal to the product of multiplication of the relative currents  Figure 2E depicts the relative peak currents upon co-application of mevastatin with xanthohumol, mevastatin applied alone, and xanthohumol applied alone. The concentration of xanthohumol was equal to 6.0 µM because such a concentration was high enough to significantly inhibit Kv1.3 channels in Jurkat T cells [20]. The relative peak current for mevastatin co-applied with xanthohumol was equal to 0.30 ± 0.11 (n = 25). This current was significantly smaller than the currents obtained for mevastatin (see above) and xanthohumol applied alone (0.72 ± 0.14, (n = 10), p < 0.05, one-way ANOVA). The relative peak current upon co-application of simvastatin with xanthohumol was equal to the value calculated for simvastatin applied alone (not shown). Since the application of xanthohumol did not change the inactivation rate of Kv1.3 currents [20], the effect of co-application of simvastatin or mevastatin with xanthohumol on the inactivation rate of the currents was not studied in detail. Figure 2F depicts relative peak currents upon co-application of mevastatin with acacetin, mevastatin applied alone, and acacetin applied alone. The concentration of acacetin was equal to 30.0 µM because such a concentration was high enough to significantly inhibit Kv1.3 channels in Jurkat T cells [19]. The concentration of the statins was previously mentioned. The relative peak current for mevastatin co-applied with acacetin was equal to 0.30 ± 0.09 (n = 31). This value was significantly smaller than the values calculated for mevastatin (see above, p < 0.05, one-way ANOVA) and acacetin applied alone (0.64 ± 0.14 (n = 19), p < 0.05, one-way ANOVA). The relative peak current upon the co-application of simvastatin with acacetin was not significantly smaller than the value for simvastatin applied alone (not shown). Figure 2G depicts relative peak currents upon co-application of simvastatin with chrysin, simvastatin applied alone, and chrysin applied alone, respectively, whereas Figure 2H shows the same data calculated upon co-application of mevastatin with chrysin, mevastatin applied alone, and chrysin applied alone. The concentration of chrysin was equal to 30.0 µM because such a concentration was high enough to significantly inhibit Kv1.3 channels in Jurkat T cells [19]. The concentration of the statins was previously mentioned. The relative peak current upon co-application of simvastatin with chrysin was equal to 0.25 ± 0.11 (n = 21). This value was significantly smaller than the values calculated for simvastatin (see above, p < 0.05, one-way ANOVA) and chrysin applied alone (0.57 ± 0.13 (n = 18), p < 0.05, one-way ANOVA). The inactivation time constant upon co-application of simvastatin with chrysin was equal to 20.64 ± 5.16 (n = 9). This value was not significantly different from the value calculated for simvastatin applied alone (p > 0.05, one-way ANOVA), but it was significantly smaller than the value obtained for chrysin applied alone (34.7 ± 15.46 (n = 9), p < 0.05, one-way ANOVA). The relative peak currents for mevastatin co-applied with chrysin were equal to 0.26 ± 0.11 (n = 21). This value was significantly smaller than the values calculated for mevastatin (see above, p < 0.05, one-way ANOVA) and chrysin applied alone (p < 0.05, one-way ANOVA). The inactivation time constant for simvastatin and mevastatin co-applied with chrysin was not significantly different from the value calculated for simvastatin and mevastatin applied alone (not shown).
The fact that the magnitude of channel inhibition upon co-application of the statins with selected flavonoids is significantly stronger than the effect exerted of each compound applied alone may be a consequence of the additivity of their inhibitory effects. If two channel inhibitors act on the same molecular target independently of each other, their inhibitory effects may be additive. In such a case, the relative peak current upon co-application of the inhibitors is equal to the product of multiplication of the relative currents upon application of each of the inhibitors alone. Table 1 shows the relative peak currents upon co-application of the statins with the selected flavonoids, calculated applying the additive inhibitory effect model, compared to the values obtained under experimental conditions. Statistical significance between theoretical and experimental values was estimated by applying Student's unpaired t-test. The difference is considered to be statistically significant when p < 0.05. The term significant (+) means that the inhibitory effect is significantly stronger than it is predicted by the additive inhibition model, whereas the term significant (−) means that it is significantly weaker. Table 1 shows that the magnitude of the inhibitory effect upon co-application of simvastatin with chrysin and mevastatin with all the compounds except for 8-prenylnaringenin was not significantly different from the magnitude predicted by the additive inhibition model. However, in the case of both statins co-applied with 8-prenylnaringenin and simvastatin co-applied with 6-prenylnaringenin, the inhibitory effect was significantly stronger than it was predicted by the additive inhibition model. On the other hand, in the case of simvastatin co-applied with xanthohumol and acacetin, the inhibitory effect was significantly weaker than predicted by the theoretical model.

Cell Viability in the Presence of Studied Compounds
Simvastatin and mevastatin exhibited a dose-dependent ability to inhibit the growth of Jurkat T cells. The concentration of mevastatin that corresponds to half of its maximal inhibitory effect (IC 50 ) was 22.5 µM. In the case of simvastatin, it was not possible to determine the IC 50 value in the studied concentration range. However, the lowest cell survival in the presence of the statin was about 60% at a concentration of 40.0 µM (Figure 3).    The ability of most flavonoids (6-prenylnaringenin, chrysin, and acacetin) to inhibit Jurkat T cells growth was shown in the work of Teisseyre et al. (2018). 6-prenylnaringenin and acacetin exhibited low potency to reduce the cells' growth (IC 50 not available), whereas IC 50 = 26.2 µM was noted for chrysin [19]. In the present studies, we also determined cell viability in the presence of 8-prenylnaringenin and xanthohumol. We found that 8-prenylnaringenin, similar to the 6-prenylated derivative, is characterized by low activity as a growth inhibitor. However, the presence of the prenyl group in the structure of xanthohumol, which has an open C-ring with a hydroxyl group on the B-ring, may contribute to increasing the ability of the compound to kill Jurkat cells (IC 50 = 32.5 µM) ( Table 2). The potency of statins, simvastatin, and mevastatin to increase the activity of flavonoids was investigated. In the presence of mevastatin, we observed a much lower value of IC50 for all studied flavonoids, except acacetin (IC 50 not available). Simvastatin and mevastatin increased cell growth inhibition by prenylated flavanones and chrysin ( Table 2). The most significant changes between the activity of pure flavonoid and its combinations with statins were observed in the case of 8-prenylnaringenin ( Figure 4A). However, the highest potency of Jurkat T cells growth inhibition was noticed for a combination of xanthohumol and mevastatin ( Figure 4B).

Apoptosis Induction
Both studied statins induced the activity of caspase-3 (p < 0,05); however, the potency of mevastatin was more significant than simvastatin ( Figure 5A). In the case of single flavonoids, only acacetin ( Figure 5B) and chrysin ( Figure 5C) increased the activity of the studied enzyme. However, the combination use of acacetin and statins did not change the effect obtained for flavonoids only ( Figure 5B). It is worth emphasizing that an increase in caspase-3 activity was noted with the simultaneous application of mevastatin with other

Apoptosis Induction
Both studied statins induced the activity of caspase-3 (p < 0,05); however, the potency of mevastatin was more significant than simvastatin ( Figure 5A). In the case of single flavonoids, only acacetin ( Figure 5B) and chrysin ( Figure 5C) increased the activity of the studied enzyme. However, the combination use of acacetin and statins did not change the effect obtained for flavonoids only ( Figure 5B). It is worth emphasizing that an increase in caspase-3 activity was noted with the simultaneous application of mevastatin with other flavonoids as opposed to their solitary use. The most significant changes were observed when mevastatin was administered in combination with 8-prenylnaringenin. The addition of simvastatin to prenylated derivatives also enhanced the induction of caspase-3 activity. However, we did not observe any statistically significant differences between the activity of this enzyme in the presence of other flavonoids and its activity in the presence of their combinations with simvastatin ( Figure 5D-F). Each column represents the mean relative activity of the enzyme normalized to the control derived from non-treated cells. The results were the means ± SDs from three independent experiments. The statistical significance was determined using Student's t-test. We compared the results obtained for each compound/combination with the results obtained from the samples containing no compounds (* p < 0.05; ** p < 0.001). We also determined the differences between results obtained for single flavonoid and its combination with statins (# p < 0.05; # p < 0.001).
Caspase-3 is one specific effector caspase, a protein that is cleaved and thus activated upon the initiation of apoptosis. Cleaved caspase-3 propagates an apoptotic signal through enzymatic activity on downstream targets. Both uncleaved and cleaved versions of the enzyme are strong indicators of cell death induction. In our study, cleaved forms of caspase-3 increased upon co-application of the drugs, which was demonstrated using the Western blot technique (Figure 6). through enzymatic activity on downstream targets. Both uncleaved and cleaved versions of the enzyme are strong indicators of cell death induction. In our study, cleaved forms of caspase-3 increased upon co-application of the drugs, which was demonstrated using the Western blot technique ( Figure 6). In our studies, the effect of the studied compound on the mitochondrial membrane potential (MMP) was determined. MMP was expressed as a fraction of cells with green fluorescence (490 nm/525 nm) in relation to the fraction of cells with red fluorescence (540 nm/590 nm), which corresponded to the monomers and aggregates' fluorescence, respectively. We found that both statins increase the fraction of JC-10 monomers. Moreover, the effect noted in the presence of mevastatin was more significant compared to simvastatin. ( Figure 7A). A reduction in MMP was also observed in the cells incubated with acacetin and chrysin. We observed that the activity of these flavonoids was potentiated by simvastatin as well as mevastatin ( Figure 7B-C). We did not find any changes between the fraction of JC-10 monomers in control cells and in the cells cultured with xanthohumol and prenylated derivatives of naringenin. The presence of simvastatin only slightly increased In our studies, the effect of the studied compound on the mitochondrial membrane potential (MMP) was determined. MMP was expressed as a fraction of cells with green fluorescence (490 nm/525 nm) in relation to the fraction of cells with red fluorescence (540 nm/590 nm), which corresponded to the monomers and aggregates' fluorescence, respectively. We found that both statins increase the fraction of JC-10 monomers. Moreover, the effect noted in the presence of mevastatin was more significant compared to simvastatin. ( Figure 7A). A reduction in MMP was also observed in the cells incubated with acacetin and chrysin. We observed that the activity of these flavonoids was potentiated by simvastatin as well as mevastatin ( Figure 7B-C). We did not find any changes between the fraction of JC-10 monomers in control cells and in the cells cultured with xanthohumol and prenylated derivatives of naringenin. The presence of simvastatin only slightly increased the ability of these flavonoids to reduce MMP. However, statistically significant changes were only noted in the case of mevastatin ( Figure 7D-F).  The results were the means ± SDs from three independent experiments. Statistical significance was determined using Student's t-test. We compared the results obtained for each compound/combination with the results obtained from the samples containing no compounds (* p < 0.05; ** p < 0.001). We also determined the differences between results obtained for single flavonoid and its combination with statins (# p < 0.05). The results were the means ± SDs from three independent experiments. Statistical significance was determined using Student's t-test. We compared the results obtained for each compound/combination with the results obtained from the samples containing no compounds (* p < 0.05; ** p < 0.001). We also determined the differences between results obtained for single flavonoid and its combination with statins (# p < 0.05).

Discussion
The results of this study showed that the co-application of simvastatin with mevastatin did not further increase the magnitude of the inhibitory effect exerted by the statins on Kv1.3 channels in human Jurkat T cells. Our results published earlier showed that both simvastatin and mevastatin inhibited Kv1.3 channels in Jurkat T cells, probably via a complex mechanism [12]. A lack of augmentation of the channel inhibition upon co-application of simvastatin and mevastatin may suggest that both statins share the mechanism of inhibition of the channels.
In contrast, co-application of both statins with most of the flavonoids caused a significant augmentation of the inhibition of Kv1.3 channels. The magnitude of inhibition upon co-application of the statins with the flavonoids could be described by applying the additive inhibition model. This model enabled calculations of relative peak currents not significantly different from experimental values in the case of simvastatin co-applied with chrysin and mevastatin co-applied with 6-prenylnaringenin, xanthohumol, acacetin, and chrysin.
However, our data also showed that inhibition upon co-application of the statins with the flavonoids might not be a result of simple additivity of inhibitory effects of the inhibitors. First of all, the inhibition upon co-application of simvastatin and mevastatin with 8-prenylnaringenin and simvastatin with 6-prenylnaringenin was significantly more potent than predicted by the additive model. On the other hand, inhibition upon co-application of simvastatin with xanthohumol and acacetin was significantly less potent than predicted by this model. In this case, no additivity was observed. The inhibition was comparable to the inhibitory effect of simvastatin applied alone. Finally, co-application of the statins with the flavonoids did not further accelerate the inactivation rate of the channels. Such an acceleration was expected if the inhibitory effects of co-applied compounds were additive. Inactivation time constants upon co-application of the statins with the flavonoids were comparable to the values obtained for the statins applied alone, except for simvastatin and mevastatin co-applied with acacetin, where they were significantly higher. Altogether, our results demonstrate that the mechanism of the channel inhibition may be more complex, including a synergistic action of both inhibitors on the channels in the case of co-application of the statins with 8-prenylnaringenin and 6-prenylnaringenin. More studies are necessary to further elucidate this problem.
Many studies have demonstrated the cytotoxic activity of statins against breast cancer [25,26], colon cancer [27], and leukemia [28]. However, only a few reports deal with the influence of combined treatment with statins and flavonoids on cancer cell survival.
In our previous studies, 6-prenylnaringenin, chrysin, and acacetin were identified as growth inhibitors of Jurkat T cells [19]. In this study, the cytotoxic effect of two additional flavonoid derivatives, 8-prenylnaringenin and xanthohumol, was also observed. In addition, both studied statins exerted a dose-dependent cytotoxic effect on the studied cancer cells. The most important result was the observation that co-treatment of Jurkat T cells with the flavonoids and the statins caused a reduction in the concentration needed to produce a 50% reduction in cell growth (IC 50 ). This may suggest that the statins may act as effective amplifiers of flavonoid activity as growth inhibitors of cancer cells. The most significant, almost 10-fold, was the reduction in IC50 in the case of xanthohumol co-applied with mevastatin. On the other hand, no significant reduction in the IC50 value was observed upon co-application of xanthohumol with simvastatin. In the case of both prenylated derivatives of naringenin and chrysin, the IC50 value upon co-application of the compounds with mevastatin was lower than the value upon the co-application with simvastatin. This may suggest that a co-application of the flavonoids with mevastatin is more effective than co-application with simvastatin. This may be due to the fact that mevastatin appeared to be a more effective inhibitor of the growth of Jurkat T cells than simvastatin.
Anticancer activity of different compounds usually results from their ability to induce apoptosis of cancer cells. Apoptosis is a stringently organized process regulated by a series of signal transduction cascades and cellular proteins [29]. Pro-apoptotic proteins are classified into Caspases and BCL-2 families. Caspases are divided into three subtypes: inflammatory caspases; initiator caspases, in intrinsic and extrinsic apoptotic pathways; and effector caspases. Depending on the type of cells, statins, as well as flavonoids, can induce different molecular pathways initiating apoptosis [30]. Apoptosis involves a disruption of mitochondrial membrane integrity that is decisive for the cell death process. Therefore, in this study, the apoptotic effects of studied compounds on Jurkat T cells were evaluated by three methods: caspase-3 activity, caspase-3 Western blot analysis, and loss of mitochondrial membrane potential (MMP). All results of the apoptotic assays indicate that Jurkat T cells went along the apoptotic pathway after mevastatin and simvastatin treatment. The potency of the mevastatin was more significant than that of simvastatin, as shown by means of an increase in caspase-3 activity and loss of MMP, in addition to the presence of caspase-3 fragments. In our previous research, we have shown that statins applied alone and together with flavonoids induce apoptosis in sensitive and resistant to doxorubicin colorectal cancer cells [15]. In this study on Jurkat T cells, statins were also co-applied with the selected flavonoids. Our results showed a correlation with studies by Watanabe et al. [31]. Similarly, we also found that inhibition of the growth of Jurkat T cells by acacetin and chrysin occurs due to the activation of caspase-3. This observation was also reflected in the reduced value of the MMP upon application of these compounds.
It is known that antiproliferative and pro-apoptotic effects of the statins on Kv1.3channel expressing cancer cells may be related to the inhibition of Kv1.3 channels [12,15]. Results of this study may indicate that ability of statins and the statins co-applied with flavonoids to inhibit cancer cell growth and to induce the apoptosis of these cells may be related to the effectiveness of inhibition of Kv1.3 channels in these cells. The ability of statins and the statins co-applied with the flavonoids, except for acacetin, to inhibit growth and induce apoptosis of Jurkat T cells correlates with their ability to inhibit Kv1.3 channels in these cells. For example, it was shown that the most significant inhibition of cancer cell growth and induction of apoptosis occurs in the case of co-application of statins with prenylated derivatives of naringenin and mevastatin co-applied with xanthohumol. This correlates with these compounds' ability to inhibit the channels upon co-application, which is the most significant in the case of the co-application of statins with prenylated derivatives of naringenin and mevastatin co-applied with xanthohumol. On the other hand, upon co-application of simvastatin and xanthohumol, no significant change in the inhibition of Kv1.3 channels was observed, and this correlates with a lack of effect of co-application on cancer cell growth and apoptosis.
As mentioned earlier, because of their ability to inhibit Kv1.3 channels and lipophilicity, statins may potentially have clinical applications in supporting the chemotherapy of some cancer types [10,12]. In the area of clinical application, a very important question is the question of the required dose. The dose should be as low as possible due to possible unwanted side effects. Reducing the required dose can be achieved by the co-application of two drugs that exert synergistic or additive effects on their molecular target when applied together. In such a case, satisfactory therapeutic results can be obtained with both drugs applied at low concentrations. A simultaneous application of statins and flavonoids may reduce the required therapeutic dose.
Another problem that has to be resolved while considering the putative application of the tested compounds in medicinal practice is the question of their bioavailability. Cellular and molecular mechanisms underlying therapeutic applications of chrysin [32], xanthohumol, acacetin [33], and 8-prenylnaringenin in various cancers have been gathered and discussed in many papers. However, the low bioavailability of flavonoids such as chrysin [34] due to rapid metabolism and excretion causes its use to be less beneficial when compared to other anticancer components from natural plants [35]. This problem may also be resolved by combination therapies, which entail the combined use of two cancer cell growth inhibitors. The simultaneous use of statins and flavonoids applied at low concentrations can increase their effectiveness and the safety of therapy.

Cell Culture and Solutions
Human leukemic T cell line, Jurkat (clone E6-1), was purchased from American Type Culture Collection (Manassas, VA, USA). The cells were grown in RPMI 1640 medium  [36,37]. The chemicals used in preparation of the solutions were purchased from the Polish Chemical Company (POCH, Gliwice, Poland), except for HEPES and EGTA, which were  [36,37]. The chemicals used in preparation of the solutions were purchased from the Polish Chemical Company (POCH, Gliwice, Poland), except for HEPES and EGTA, which were purchased from Sigma-Aldrich. The examined compounds (statins and flavonoids) were purchased from Alexis Biochemicals (Lausen, Switzerland).

Patch-Clamp Recordings
Dishes with examined Jurkat T cells were placed under an inverted Olympus IMT-2 microscope. Washing solutions and solutions containing examined statins and flavonoids were applied using eight-channel gravitation perfusion system (ALA Scientific Instruments, Farmingdale, NY, USA). Pipettes were pulled from a borosilicate glass (Hilgenberg, Germany). The pipette resistance was higher than 3.0 MΩ. Kv1.3 currents in Jurkat T cells were recorded applying the whole-cell patch-clamp technique [38]. The currents were recorded using an EPC-7 Amplifier (HEKA, Germany), low-pass filtered at 3.0 kHz, and digitised using a CED Micro 1401 analog-to-digital converter (Cambridge, UK) with a sampling rate of 10.0 kHz. The influence of selected compounds on the activity of the channels was studied by applying the voltage ramp protocol. During application of a voltage ramp, the membrane of examined cell was gradually depolarised from −100.0 mV up to +40.0 mV. The ramp duration was 340.0 ms. The ramps were applied every 30.0 s, and the holding membrane potential kept at the meantime was −90.0 mV [12]. Upon application of the voltage ramp protocol, potassium currents in Jurkat T cells could be stably recorded for at least 20 min after "break-in" to the whole-cell configuration. During the off-line analysis, the leak current estimated at +40.0 mV was subtracted from the total ramp current recorded at this voltage, giving the value of peak Kv1.3 currents.
In order to study the influence of selected compounds on the channel inactivation kinetics in more detail, another protocol containing a sequence of depolarizing voltage steps from the holding potential of −90.0 mV to +60.0 mV (500.0 ms step duration) was applied every 30.0 s [12]. All experiments were carried out at room temperature (22-24 • C). The data are presented as mean ± standard deviation.

MTT Assay
The influence of studied compounds on the viability of Jurkat T cells was determined using MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide) test as described previously [19]. The impact of single statins or flavonoids was examined in the concentration range of 2.5-40.0 µM. In case of compounds' combinations, flavonoids were applied in the concentration range of 2.5-40.0 µM, whereas the concentration of statin (simvastatin or mevastatin) was constant (6.0 µM).

Caspase-3 Activity
The activity of caspase-3 was determined with use of Caspase-3 Colorimetric Assay Kit (GenScript), according to the manufacturer's protocol with minor modifications. In this experiment, chromophore p-nitroaniline (pNA) released by the enzyme from the labeled substrate DEVD-pNA was detected by spectrophotometric method. Jurkat T cells (3.0 × 10 6 ) were cultured in the presence of compound for 48.0 h. The cultures were maintained in humidified atmosphere of 5% CO 2 at 37 • C. In the next step, cells were centrifuged (900.0 rpm/RT/5 min); pellet in each of probes was washed with PBS and centrifuged once again (900.0 rpm/RT/5 min). Next, supernatants were removed, and the cells were re-suspended in Lysis Buffer. The samples were incubated for 60.0 min on ice and mixed every 15.0 min. Then, mixtures were centrifuged (7000.0 rpm/4 • C/1 min) and concentration of proteins in cytosolic extract (supernatant) was determined using Bradford method.

MMP Investigation
In order to determine the loss of mitochondrial membrane potential (MMP) Mitochondrial Membrane Potential Kit (Sigma-Aldrich, St. Louis, MI, USA) was used. The test is based on the fluorimetric detection of cationic, lipophilic JC-10 dye, which in nor-mal cells concentrates in the mitochondrial matrix in the form of aggregates characterized by red fluorescence (λ ex = 540/λ em = 590 nm). In apoptotic or necrotic cells, the loss of MMP leads to decomposition of aggregates to form green fluorescent monomers (λ ex = 490/λ em = 525 nm). The procedure was performed according to the manufacturer's protocol with minor modifications. In our experiments, Jurkat cells were seeded at a density of 3 × 10 4 onto 96-well and treated with examined compounds. The cultures were maintained in humidified atmosphere of 5% CO 2 at 37 • C. Time of incubation was 48 h. In the next step, plates were centrifuged (500.0 rpm/RT/5 min), and the medium was removed. Then, 100.0 µL of Dye Loading Solution was added to each well, and the plates were incubated 5% CO 2 at 37 • C for 30.0 min. In the final step, 50.0 µL of Assay Buffer B was added, and the plates were centrifuged (800 rpm/RT/2 min). The plates were incubated for 20.0 min, and the fluorescence intensities (ex/em = 485/525 nm and ex/em = 540/590 nm) were measured with use of Tecan Infinite 200 PRO plate reader.

Western Blot Analysis
Equal amounts of proteins after isolation were separated by 7% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), after which the resolved proteins were transferred to a nitrocellulose membrane (Whatman GmbH, Dassel, Germany). The membrane was blocked with 5% milk, followed by exposure to the appropriate antibodies (caspase-3 Antibody Novus 31A11067). β-actin antibody was purchased from Sigma-Aldrich. All bands were visualized using horseradish peroxidase-conjugated secondary antibodies. Secondary antibodies were supplied by Santa Cruz Biotechnology.

Data Analysis
The magnitude of the channel inhibition was calculated as the relative current recorded upon application of the studied compound. The current was defined as I/Icontr; where: I-the peak Kv1.3 current upon an application of an examined compound, Icontr-the peak current recorded on the same cell under control conditions. Inactivation kinetics were fitted by the single exponential function and described by the value of inactivation time constant. In all experiments, statistical analysis was performed by applying the Student's unpaired t-test or one-way ANOVA test. The corrected Bonferroni p values were taken into account while performing ANOVA tests. The results were considered statistically significant when p < 0.05.

Conclusions
Simvastatin, mevastatin, and some flavonoids inhibit the activity of Kv1.3 channels in Jurkat T cells. This effect may correlate with the ability of these compounds to induce apoptosis through a reduction in MMP in studied cancer cells. Moreover, the statins may potentiate the biological activity of selected flavonoids. The simultaneous application of these types of chemical compounds could be a promising anticancer strategy. It is suggested that the usage of compounds combination allows for a reduction in the effective dose of therapeutics and thus for a reduction in detrimental side effects in patients.